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HomeScience SignalingVol. 14, No. 705The injury response to DNA
damage in live tumor cells promotes antitumor immunity
Back To Vol. 14, No. 705
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Research ArticleCANCER THERAPY
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The injury response to DNA damage in live tumor cells promotes
antitumor immunity

Ganapathy Sriram https://orcid.org/0000-0002-1651-3607, Lauren E. 
Milling https://orcid.org/0000-0002-1053-0887, Jung-Kuei Chen https:/
/orcid.org/0000-0003-4964-2031, Yi Wen Kong https://orcid.org/
0000-0002-4223-971X, Brian A. Joughin https://orcid.org/
0000-0003-1022-9450, Wuhbet Abraham, Susanne Swartwout, Erika D. 
Handly https://orcid.org/0000-0001-8788-3050, Darrell J. Irvine
https://orcid.org/0000-0002-8637-1405 [email protected], and Michael
B. Yaffe https://orcid.org/0000-0002-9547-3251 [email protected]
Science Signaling * 19 Oct 2021 * Vol 14, Issue 705 * DOI: 10.1126/
scisignal.abc4764
 

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  * Tailoring cancer vaccines to patients
  * Abstract
  * INTRODUCTION
  * RESULTS
  * DISCUSSION
  * MATERIALS AND METHODS
  * Acknowledgments
  * Supplementary Materials
  * REFERENCES AND NOTES

Tailoring cancer vaccines to patients

Immunotherapies boost the body's immune system to detect and kill
cancer cells. The immune response can be enhanced by molecules
released from dying tumor cells, leading to proposals to use dead
tumor cells as adjuvants in immunotherapeutics and cancer vaccines.
However, Sriram et al. found that, in some cases, it is the remaining
live tumor cells that are immunogenic. Treating cultured tumor cells
with select DNA-damaging chemotherapies (and specific doses thereof)
stimulated cytotoxic T cell activation when the live, not dead, cell
fraction was injected back into the tumor in mice. Combining these
cells with clinically used immune checkpoint inhibitors resulted in
tumor regression and antitumor immune memory. The findings
potentially identify a patient-specific approach to making
immunotherapies more durably and broadly effective.

Abstract

Although immune checkpoint blockade (ICB) has strong clinical benefit
for treating some tumor types, it fails in others, indicating a need
for additional modalities to enhance the ICB effect. Here, we
identified one such modality by using DNA damage to create a live,
injured tumor cell adjuvant. Using an optimized ex vivo coculture
system, we found that treating tumor cells with specific
concentrations of etoposide, mitoxantrone, or doxorubicin markedly
enhanced dendritic cell-mediated T cell activation. These
immune-enhancing effects of DNA damage did not correlate with
immunogenic cell death markers or with the extent of apoptosis or
necroptosis; instead, these effects were mediated by live injured
cells with activation of the DNA-PK, ATR, NF-kB, p38 MAPK, and RIPK1
signaling pathways. In mice, intratumoral injection of ex vivo
etoposide-treated tumor cells in combination with systemic ICB (by
anti-PD-1 and anti-CTLA4 antibodies) increased the number of
intratumoral CD103^+ dendritic cells and circulating
tumor-antigen-specific CD8^+ T cells, decreased tumor growth, and
improved survival. These effects were absent in Batf3^-/- mice and in
mice in which the DNA-damaging drug was injected directly into the
tumor, due to DNA damage in the immune cells. The combination
treatment induced complete tumor regression in a subset of mice that
were then able to reject tumor rechallenge, indicating that the
injured cell adjuvant treatment induced durable antitumor
immunological memory. These results provide a strategy for enhancing
the efficacy of immune checkpoint inhibition in tumor types that do
not respond to this treatment modality by itself.

INTRODUCTION

Cytotoxic chemotherapy remains a mainstay of cancer treatment. It is
estimated that >60% of all patients with cancer will receive some
type of initial treatment beyond surgery that includes DNA damaging
drugs and/or antimicrotubule agents, particularly in patients with
more advanced stage disease (1). These conventional chemotherapy
treatments result in extended survival and/or cure, depending on the
specific tumor type; however, toxic side effects and the development
of resistance and subsequent tumor relapse are frequently seen.
Therapeutic manipulation of the immune system has now emerged as an
alternative approach to anticancer therapy, as a consequence of the
development of inhibitors targeting the immune checkpoint proteins
programmed cell death protein-1 (PD-1), programmed death ligand-1
(PD-L1), and cytotoxic T-lymphocyte associated protein 4 (CTLA4) (2).
Certain tumor types show impressive clinical responses to these
agents, particularly melanoma (3), non-small cell lung cancer (NSCLC)
(4, 5), and microsatellite instability-high colon cancer (6, 7).
However, most patients with many common tumor types, including breast
cancer (8, 9), ovarian cancer (10, 11), and microsatellite-stable
colon cancer (12), show much lower response rates. It has been
estimated that the overall percentage of all patients with cancer who
will respond to immune checkpoint inhibitors alone is less than 13% (
13).
For many tumor types, immunotherapy has been reserved as a second- or
third-line treatment option in patients who have failed prior
treatment with cytotoxic agents (14). However, early combination
treatment using chemotherapy and immune checkpoint inhibitors as a
first line therapeutic modality was recently approved for advanced
NSCLC that lack epidermal growth factor or anaplastic lymphoma kinase
mutations using cisplatin and pembrolizumab (15), for head and neck
squamous cell carcinomas (HNSCCs) using platinum agents,
5-fluorouracil (5-FU), and pembrolizumab (16), and for
triple-negative breast cancer (TNBC) using pembrolizumab and either
paclitaxel or gemcitabine plus carboplatin (17).
Data supporting this approach come from the KEYNOTE-189 trial, which
showed a median progression-free survival of 8.8 months in patients
with NSCLC that were treated with a combination of cisplatin or
carboplatin, pemetrexed, and pembrolizumab, compared to 4.9 months in
patients who were treated with chemotherapy alone. However, over 65%
of the patients who received this chemotherapy and immunotherapy
combination continued to have progressive disease (15). Similarly,
the KEYNOTE-048 trial, performed in patients with recurrent
unresectable HNSCC in which the tumor contained more than 1% of cells
staining positively for PD-L1, failed to show any improvement in
progression-free survival in patients treated with cisplatin or
carboplatin, 5-FU, and pembrolizumab compared to those treated with
the same chemotherapy plus cetuximab, although there was an increase
in median overall survival from 10.7 to 13 months when pembrolizumab
was included in the combination (16). Last, in the KEYNOTE-355 trial
involving patients with locally recurrent but inoperable or
metastatic TNBC, the median progression-free survival at 12-months in
patients whose tumors showed >=10% PD-L1-positive staining increased
from 23 to 39% by the addition of immunotherapy to chemotherapy
compared with chemotherapy alone, but more than 70% of the
intention-to-treat population, had progressive disease by this time (
17). Identifying mechanisms that would enhance response rates to the
combination of immune checkpoint blockade (ICB) and chemotherapy--and
prolong the durability of the response--remains an unmet clinical
need.
On the basis of our long-standing interest in cross-talk between the
DNA damage response and signaling pathways that mediate tumor cell
survival, apoptotic cell death, and innate immune activation (18-23),
we investigated whether signaling pathways activated in response to
specific types of DNA-damaging chemotherapy could enhance subsequent
antitumor immune responses. Although the ability of specific
chemotherapeutic compounds to enhance cross-presentation of tumor
antigens by dendritic cells (DCs) has been characterized as
"immunogenic cell death" (24-27), we found that chemotherapy-induced
cell stress signaling in live injured cells, but not the presence of
dead cells, was the primary determinant of T cell immunity. This
effect is mediated by DNA-dependent protein kinase (DNA-PK), ataxia
telangiectasia and Rad3-related kinase (ATR), nuclear factor kB
(NF-kB), p38 mitogen-activated protein kinase (p38 MAPK), and
receptor-interacting Ser/Thr kinase 1 (RIPK1) signaling in the
injured tumor cells, depending on the drug and cell type.
Furthermore, we show that direct intratumoral injection of ex vivo
chemotherapy-treated cells as an injured cell adjuvant, in
combination with systemic immune checkpoint blockade, drives
antitumor immunity and tumor regression in murine melanoma models, in
contrast to ICB alone.

RESULTS

Etoposide- and mitoxantrone-treated tumor cells can induce
DC-mediated OT-1 T cell interferon-g responses in vitro

To identify how tumor cell stress and injury after DNA-damaging
chemotherapy could potentially influence antitumor immune function,
we treated B16F10 melanoma tumor cells expressing ovalbumin (B16-Ova)
or MC-38 colon cancer cells expressing ovalbumin (MC-38-Ova; fig.
S1A) with various doses of the common clinically used
chemotherapeutic agents doxorubicin, etoposide, mitoxantrone,
cisplatin, oxaliplatin, cyclophosphamide, irinotecan, camptothecin,
paclitaxel, or 5-FU and examined them for immunogenicity by assaying
for their ability to induce DC-mediated interferon-g (IFN-g)
responses in CD8^+ T cells. After tumor cell treatment with
chemotherapeutic drugs for 24 hours, the drugs were washed out with
two or more changes of medium, and the treated cell mixture
(including both injured and dead cells) was then cocultured with
primary bone marrow-derived DCs (BMDCs) for an additional 24 hours (
Fig. 1A). Purified naive CD8^+ T cells obtained from the spleens of
OT-1 mice were then added to the drug-treated B16-Ova cells/BMDC
coculture, and the appearance of IFN-g^+ CD8^+ T cells was quantified
12 to 15 hours later by intracellular staining and flow cytometry (
Fig. 1B). CD8^+ T cells of OT-1 mice express a transgenic T cell
receptor recognizing ovalbumin residues 257 to 264 in the context of
H2-K^b (28, 29).
[scisignal]
Fig. 1. An experimental system to assess DC-mediated T cell IFN-g
responses to genotoxically damaged tumor cells.
(A) Schematic of the in vitro experimental system. B16-Ova or
MC-38-Ova cells were treated with DNA-damaging agents for 24 hours,
washed, and incubated with primary bone marrow-derived dendritic
cells (BMDCs) for another 24 hours. After this, OT-1 CD8^+ T cells
expressing a T cell receptor transgene that specifically recognizes
the Ova-derived peptide SIINFEKL in the context of H2-K^b (OT-1) (28,
29) were added and evaluated for intracellular IFN-g 15 hours later.
(B) T cells resulting from the experimental protocol described in (A)
were identified by CD3 staining and re-gated for IFN-g and CD8
expression. Representative flow cytometry plots assessing IFN-g^+CD8^
+ T cell populations (boxed region) induction by treatment of B16-Ova
cells with 10 or 50 mM doxorubicin, etoposide, or mitoxantrone. (C
and D) Quantification of BMDC-mediated induction of IFN-g^+CD8^+ T
cells from five (C) or three (D) independent experiments with B16-Ova
and MC-38-Ova cells, respectively, described in (A) and (B). The
first lane marked "-" indicates the percentage of IFN-g^+CD8^+ T
cells produced by coculture of BMDCs and T cells in the absence of
tumor cells. (E) Schematic of the in vitro experimental system using
TC-1 cells treated cocultured with BMDCs as described in (A) and then
cocultured with CD8^+ T cells isolated from mice vaccinated (primed
and boosted) with an HPV-E7 peptide. As in (A), T cells were then
assessed for intracellular IFN-g 15 hours later. (F) Quantification
of IFN-g^+ CD8^+ T cells from three experiments described in (E). "-"
as described for (C) and (D). For all the above panels, error bars
indicate SEM. *P < 0.05, ***P < 0.005, and ****P < 0.0001 by ANOVA,
followed by Sidak's multiple comparisons test.
Treatment of B16-Ova or MC-38-Ova cells with either etoposide or
mitoxantrone was the most effective at inducing DC-mediated IFN-g in
OT-1 CD8^+ T cells when the treated cells were cocultured with BMDCs
(Fig. 1, C and D, and fig. S1B). The effectiveness of these
DNA-damaging drugs at inducing T cell IFN-g responses was dose
dependent for each cell line. Unexpectedly, in contrast to etoposide
and mitoxantrone (both DNA topoisomerase II inhibitors), doxorubicin
(also a topoisomerase II inhibitor but in a different drug class,
anthracyclenes) was ineffective at inducing DC-mediated IFN-g in T
cells when administered at comparable doses, despite causing similar
or higher levels of total cell death (Fig. 1, C and D, and fig. S1, C
and D). The effects of 50 mM etoposide, 10 mM mitoxantrone, and 10 mM
doxorubicin on T cell stimulation assayed by IFN-g production were
consistent with the effects of these drugs on T cell proliferation,
as assayed by carboxyfluorescein diacetate succinimidyl ester (CFSE)
dilution (fig. S1E).
To examine whether the antitumor immune response induced by
chemotherapy treatment required the artificial expression of
ovalbumin and the OT-1 T cell system, we used a second tumor cell
line, TC-1, which was previously established from mouse lung tumors
by immortalization with human papilloma virus (HPV) protein E7 (30).
To generate T cells specific for the E7 antigen, we vaccinated mice
with three doses of an HPV-E7 peptide and isolated total splenic CD8^
+ T cells 6 days after the third immunization. This bulk population
of splenic T cells was then used in drug treatment experiments with
TC-1 cells, as described above (for Fig. 1E). Similar to what was
seen in the MC-38-Ova/OT-1 experiments, TC-1 cells treated with the
indicated doses of etoposide and mitoxantrone, but not with similar
doses of doxorubicin, showed an enhanced IFN-g response (Fig. 1F),
with the strongest response to mitoxantrone at 50 mM.

Conventional markers of immunogenic cell death do not fully explain
the T cell response to mitoxantrone and etoposide treatment

In our in vitro assay system, both mitoxantrone and etoposide were
found to induce DC-mediated T cell IFN-g responses, in contrast to
doxorubicin (at 10 and 50 mM). Obeid et al. (24) reported that
mitoxantrone chemotherapy induced strong immunogenic cancer cell
death in CT26 mouse colon carcinoma cells, based on the drug's
ability to induce calreticulin exposure on the cell surface.
Prophylactic subcutaneous injection of these dying, drug-treated
cells into one flank enabled mice to resist a subsequent tumor
challenge into the opposite flank (24). In addition to externalized
calreticulin, release of high-mobility group box protein 1 (HMGB1)
and adenosine triphosphate (ATP) was also reported to serve as
canonical markers of immunogenic cell death (27).
To directly test whether these markers correlated with T cell priming
in our system, we treated B16-Ova cells with mitoxantrone, etoposide,
or doxorubicin and measured calreticulin exposure on the cell surface
at 24 hours, when the treated cells were cultured with BMDCs. We also
analyzed HMGB1 and ATP release during the first 24 hours of
chemotherapy treatment and during the 24- to 48-hour posttreatment
window, which corresponds to the full period of BMDC coculture (Fig.
2A). In previous reports, etoposide was not considered an immunogenic
cell death-inducing drug due to its inability to induce endoplasmic
reticulum (ER) stress and calreticulin exposure in CT26 cells (24),
despite inducing the release of HMGB1 and ATP (31). However, we
included etoposide in these experiments because it induced equivalent
levels of IFN-g^+CD8^+ T cells as mitoxantrone in our in vitro assay
for DC-mediated T cell responses. Doxorubicin was specifically chosen
as the third drug for comparison because, similar to etoposide and
mitoxantrone, it inhibits topoisomerase II but did not induce T cell
IFN-g responses when used at similar doses (Fig. 1, B and C),
although doxorubicin has been reported to induce calreticulin
exposure and the release of HMGB1 and ATP in CT26 cells (24, 31).
[scisignal]
Fig. 2. T cell IFN-g responses by DNA-damaged tumor cells involves
calreticulin and signaling through NF-kB, p38 MAPK, and RIPK1.
(A) Left: Percentage of B16-Ova tumor cells displaying surface
calreticulin 24 hours after the indicated treatment from a
representative of two experiments. Results from an additional
representative experiment using a second antibody are shown in fig.
S2A. Middle and right: Levels of HMGB1 and ATP in the culture medium
measured from 24 to 48 hours after the indicated treatment. Results
are from four independent experiments, with error bars indicating
SEM. *P < 0.05, **P < 0.01, ***P < 0.005, and ****P < 0.0001 compared
to DMSO-treated control by ANOVA, followed by Dunnett's multiple
comparisons test. (B) Schematic of the siRNA experiment testing the
role of B16-Ova cell calreticulin in BMDC-mediated IFN-g induction in
T cells. (C) Left: Quantification of the fold change in the
percentage of IFN-g^+CD8^+ T cells from the experiment in (B). Data
for each calreticulin knockdown condition are normalized to the
respective drug-treated control knockdown condition. Results
represent three independent experiments with error bars indicating
SEM. Data were analyzed by ANOVA followed by Sidak's multiple
comparisons test comparing drug-treated knockdown to drug-treated
controls. ****P < 0.0001. Right: Quantification of calreticulin
knockdown efficiency by Western blotting (WB). First lane (-) is
defined as in Fig. 1C. (D to G) Quantification of the fold change in
the percentage of IFN-g^+CD8^+ T cells induced by BMDCs after
incubation with etoposide- or mitoxantrone-treated B16-Ova (D and E)
or MC-38-Ova (F and G) cells that were cotreated with the indicated
DNA-damaging agent plus Z-VAD, necrostatin-1 (Nec-1), Bay 11-7085
(inhibitor of NF-kB signaling), or SB202190 (p38 MAPK inhibitor).
First lane (-) defined as in Fig. 1C. Data are normalized to the
condition in which B16-Ova cells are treated with etoposide alone or
mitoxantrone alone, respectively. Results from three independent
experiments with error bars indicating SEM. *P < 0.05 and ****P <
0.0001 wherein each cotreatment condition (etoposide or mitoxantrone
+ Z-VAD, Nec-1, NF-kBi, or p38i) is compared with the etoposide or
mitoxantrone alone condition using ANOVA, followed by Dunnett's
multiple comparisons test.
Using two calreticulin antibodies, all three drugs elicited only low
levels of calreticulin exposure at this time point (24 hours), with
<20% of the cells staining positively (Fig. 2A and fig. S2A, left).
Mitoxantrone treatment induced the highest percentage of cells with
externalized calreticulin when analyzed by flow cytometry after 24
hours of drug exposure. Treatment with either low or high etoposide
concentrations caused an intermediate percentage of cells to display
calreticulin surface exposure (<10%), whereas doxorubicin treatment
resulted in the lowest percentage of cells with calreticulin exposure
(<5%).
There was no correlation between HMGB1 or ATP release and induction
of immunogenicity as measured by IFN-g production in stimulated T
cells (Fig. 2A, middle and right). Cells treated with etoposide
showed the lowest levels of HMGB1 release into the medium during the
24- to 48-hour posttreatment window (i.e., when the tumor cells were
exposed to BMDCs; Fig. 2A, middle), despite being highly immunogenic.
In contrast, but similar to what was observed with mitoxantrone (10
mM) treatment, doxorubicin treatment led to high levels of HMGB1
release, despite its inability to promote BMDC-mediated T cell IFN-g
responses at this dose. Similar trends in HMGB1 release were observed
during the first 24 hours of treatment (fig. S2B). During the first
24 hours of drug exposure, ATP release from B16-Ova cells after
doxorubicin and mitoxantrone treatment but not etoposide treatment
was higher than the control (fig. S2C), despite the observation that
only mitoxantrone- and etoposide-treated B16-Ova cells induced
DC-mediated IFN-g in T cells. During the 24- to 48-hour window after
drug exposure, when B16-Ova cells are cocultured with BMDCs, the
release of ATP by B16-Ova cells for all drug/dose treatment
conditions was comparable to the baseline levels induced by the
vehicle control condition (Fig. 2A, right). Overall, whereas the
calreticulin exposure on the total treated cell mixture partially
explains the differences in chemotherapy-induced tumor cell
immunogenicity, neither HMGB1 release nor ATP secretion was
predictive of DC-mediated T cell IFN-g responses for all three
chemotherapy agents.
Given the lack of correlation between HMGB1 or ATP release from the
drug-treated tumor cells and dose-specific induction of IFN-g in T
cells, we focused on the specific role of calreticulin. To directly
measure the contribution of calreticulin to DC-mediated T cell IFN-g
responses in our assay, we repeated the experiments outlined above
(in Fig. 1A) after knockdown of calreticulin in B16-Ova cells using
targeted small interfering RNA (siRNA) (Fig. 2B). Knockdown of
calreticulin before drug treatment reduced the percentage of
DC-mediated IFN-g^+ T cells after mitoxantrone exposure by ~80% and
the percentage of DC-mediated IFN-g^+ T cells after etoposide
exposure by ~50% (Fig. 2C). These data suggest that in B16-Ova cells,
calreticulin can account for a substantial amount of the drug-induced
antitumor immunogenicity, consistent with its previously described
role in immunogenic cell death (24). However, two other previously
described canonical markers of immunogenic cell death, HMGB1 and ATP,
failed to explain the observed results.

T cell IFN-g responses by BMDCs cocultured with DNA-damaged tumor
cells depend on RIPK1, NF-kB, and p38 MAPK signaling in the tumor
cells

To further explore the signaling connection between antitumor
immunogenicity and cell death, we inhibited the kinase RIPK1, which
is a known determinant of necroptosis (32); caspases, which are known
determinants of apoptosis and pyroptosis (33); the transcription
factor NF-kB, which is a critical regulatory node for survival and
cytokine production (34); and the kinase p38 MAPK, which is a
well-known master regulator of stress signaling, including signaling
downstream of DNA damage (35), all of which were activated by
etoposide and/or mitoxantrone treatment (fig. S2, D to G). B16-Ova
cells were cotreated with etoposide or mitoxantrone in combination
with the RIPK1 inhibitor necrostatin-1 (Nec-1), the pan-caspase
inhibitor Z-VAD, the NF-kB signaling inhibitor Bay 11-7085 (36), or
the p38 MAPK inhibitor SB202190 (37), and the drugs were washed out
before coculture with BMDCs. Cotreatment with Nec-1 inhibited the
ability of both etoposide- and mitoxantrone-treated B16-Ova cells,
cocultured with BMDCs, to induce IFN-g in T cells (Fig. 2, D and E),
suggesting that the ability of both etoposide and mitoxantrone to
induce immunogenicity in this model is RIPK1 dependent. In contrast,
cotreatment of B16-Ova cells with Z-VAD (validated in fig. S2D) only
marginally reduced T cell IFN-g responses (by ~12%) with etoposide
and had no effect with mitoxantrone, indicating that the process for
both agents was largely independent of caspases. Furthermore,
cotreatment of B16-Ova cells with the NF-kB signaling inhibitor Bay
11-7085 (validated in fig. S2E) and etoposide reduced the frequency
of IFN-g^+ T cells by >90%, whereas cotreatment with Bay 11-7085 and
mitoxantrone reduced the frequency of IFN-g^+ T cells by >50%,
suggesting that NF-kB signaling in both etoposide- and
mitoxantrone-treated B16-Ova cells is important for the induction of
DC-mediated T cell IFN-g responses. Last, cotreatment of B16-Ova
cells with the p38 MAPK inhibitor SB202190 and etoposide reduced the
frequency of IFN-g^+ T cells by ~22%, whereas cotreatment with
SB202190 and mitoxantrone nearly abrogated the induction of IFN-g^+ T
cells altogether. Consistent with these results, both etoposide and
mitoxantrone (which induced DC-mediated T cell IFN-g responses) but
not doxorubicin (which did not) were found to induce RIPK1 activation
in B16-Ova cells, as shown by Western blotting with an antibody to
Ser^166-phoshorylated RIPK1 (fig. S2F). Western blotting of cell
lysates with an antibody to phosphorylated p38 demonstrated p38 MAPK
activation by etoposide and mitoxantrone, as well as by doxorubicin,
suggesting that induction of p38 MAPK signaling in tumor cells is
necessary but not sufficient for the induction of IFN-g in T cells
(fig. S2G). Together, these data suggest that active signaling
through the RIPK1, NF-kB, and p38 MAPK signaling pathways after
chemotherapy treatment is involved in the induction of DC-mediated T
cell IFN-g responses. Similar results were obtained in MC-38-Ova
cells for all but RIPK1, the inhibition of which, in
etoposide-treated tumor cells, had minimal effects on T cell IFN-g
production and actually enhanced its production in response to
mitoxantrone-treated tumor cells (Fig. 2, F and G).

In situ treatment of B16-Ova tumors with direct intratumoral
etoposide does not synergize with systemic checkpoint blockade,
consistent with immune cell damage

Given the wide clinical use of etoposide in oncology (38) and the
ability of etoposide-treated B16-Ova cells to induce DC-mediated T
cell IFN-g responses ex vivo, as presented above (Fig. 1), we
reasoned that intratumoral administration of etoposide could enhance
DC function in vivo by increasing the immunogenicity of B16-Ova
cells. This would be expected to induce antigen-specific T cell
expansion in vivo, particularly if used in combination with systemic
ICB. To test this, we treated mice bearing flank B16-Ova tumors by
intratumoral administration of either saline or etoposide (three
weekly doses) in the presence or absence of systemic anti-PD1 and
anti-CTLA4 antibodies (two doses a week for three weeks) to confer
ICB (Fig. 3A). Intratumoral injection of etoposide alone had no
effect on tumor growth (Fig. 3B, top). Systemic administration of ICB
in combination with intratumoral chemotherapy also did not
significantly enhance survival beyond that seen with ICB alone (Fig.
3, B and C). Furthermore, when we examined the frequency of
circulating H2-K^b/SIINFEKL-specific CD8^+ T cells, we were unable to
detect an expansion of these cells when compared to the group that
received checkpoint blockade alone (Fig. 3D).
[scisignal]
Fig. 3. Direct intratumoral administration of etoposide does not
synergize with ICB.
(A) Schematic of the experimental design and dosing regimen used for
testing intratumoral administration of etoposide in the presence or
absence of systemic anti-PD1 and anti-CTLA4. (B) Tumor growth curves
in mice bearing B16-Ova tumors treated with intratumoral saline
(Saline IT; gray arrowheads) or etoposide (Etop IT; brown arrowheads)
in the presence or absence of systemic anti-PD1 and anti-CTLA4 (green
arrowheads). The number of mice in each group is indicated. One mouse
in the Etop IT + anti-PD1/CTLA4 group did not show tumor growth
beyond 4 mm^2 throughout the experiment and was excluded. (C)
Kaplan-Meier survival curves of the experiment in (B), compared by
log-rank test. (D) Frequency of circulating H2-K^b/SIINFEKL-specific
CD8^+ T cells from mice treated with the conditions indicated. Error
bars represent SEM. The frequency of H2-K^b/SIINFEKL tetramer-stained
CD8^+ T cells in the Etop IT + anti-PD1/CTLA4 treatment group was not
significantly different from that of the Saline IT + anti-PD1/CTLA4
group (one-tailed t test, P = 0.4553 indicated by ns). (E) Schematic
of the experiment examining etoposide cotreatment of BMDCs, T cells,
and B16-Ova cells. (F) Quantification of IFN-g^+ CD8^+ T cells (from
three independent experiments) induced by BMDCs after coculture with
etoposide-treated B16-Ova cells when both BMDCs and T cells were
exposed to etoposide compared to when only B16-Ova cells were
exposed. Error bars represent SEM. ****P < 0.0001 by ANOVA followed
by Sidak's multiple comparisons test.
However, intratumoral administration of etoposide exposes both tumor
cells and nontumor cell types such as intratumoral DCs to this
cytotoxic drug, which could potentially limit DC activation and
impair the expansion of tumor antigen-specific T cells. To test this
hypothesis, we revised the assay described above (shown in Fig. 1A)
to now include coexposure of tumor cells and either BMDCs or both
BMDCs and T cells to etoposide before the addition of OT-1 T cells (
Fig. 3E and fig. S3A). Coexposure of both BMDCs and tumor cells to
etoposide significantly reduced the appearance of IFN-g^+CD8^+ T
cells compared to exposure of B16-Ova cells alone (fig. S3A),
indicating that exposure of DCs to etoposide impairs their ability to
induce T cell IFN-g responses. Consistent with this idea, the
viability of BMDCs was significantly reduced upon exposure to
etoposide (fig. S3B). We further modified the assay to include
exposure of all of the relevant cell types--tumor cells, BMDCs, and T
cells--to etoposide, mirroring what might occur after intratumoral
injection of the drug in vivo. This triple coexposure resulted in an
even more profound loss of IFN-g^+ T cells to less than 10% of the
level seen when etoposide exposure is limited to the tumor cells
alone (Fig. 3F).

Intratumoral injection of ex vivo etoposide-treated tumor cells
synergizes with ICB to enhance the antitumor response and survival

Exposure of BMDCs and T cells to etoposide reduced the induction of
IFN-g^+CD8^+ T cells by drug-treated B16-Ova cells compared to
etoposide exposure of B16-Ova cells alone. We therefore reasoned that
the intratumoral injection of ex vivo etoposide-treated B16-Ova cells
into B16-Ova tumors in vivo, rather than intratumoral injection of
the free drug, would minimize exposure of other immune cell types in
the tumor and draining lymph node to the cytotoxic effects of
etoposide. To test this, mice bearing flank B16-Ova tumors received
intratumoral injection of either saline or ex vivo etoposide-treated
B16-Ova cells in the presence or absence of systemic checkpoint
blockade (Fig. 4A). Intratumoral administration of ex vivo
etoposide-treated tumor cells alone had no effect on subsequent tumor
progression (Fig. 4, B to D). However, when used in combination with
systemic checkpoint blockade, the mice displayed superior tumor
control compared to those that received checkpoint blockade alone,
resulting in complete tumor regressions in ~35% of mice. Furthermore,
survival was also markedly enhanced in this group (Fig. 4C). Analysis
of circulating lymphocytes in these animals revealed an enhanced
frequency of H2-K^b/SIINFEKL-specific CD8^+ T cells (Fig. 4E),
indicating that intratumoral administration of ex vivo
etoposide-treated tumor cells functions as an effective injured cell
adjuvant, which, in combination with ICB, promotes efficient T cell
priming and antitumor immunity.
[scisignal]
Fig. 4. Intratumoral administration of ex vivo DNA-damaged tumor
cells plus systemic ICB expands CD8^+ T cells and increases tumor
control and survival.
(A) Schematic of the experimental design and dosing regimen used for
testing intratumoral administration of etoposide-treated B16-Ova
cells (injured cell adjuvant) in the presence or absence of systemic
anti-PD1 and anti-CTLA4. (B) Tumor growth curves for mice treated
with intratumoral saline (Saline IT; gray arrowheads) or ex vivo
etoposide-treated B16-Ova cells (injured cell adjuvant IT; brown
arrowheads) in the presence or absence of systemic anti-PD1 and
anti-CTLA4 (green arrowheads). "n" indicates the number of mice in
each group. (C) Kaplan-Meier survival curves of the experiment in
(B). *P < 0.05 by log-rank test compared to the Saline IT + anti-PD1/
CTLA4 group. ns, P > 0.05. (D) Average tumor cross-sectional area on
day 21 for each treatment group. Error bars indicate SEM. *P < 0.05
and (ns) P > 0.05 by one-tailed t tests. (E) Frequency of circulating
H2-K^b/SIINFEKL-specific CD8^+ T cells from mice after the indicated
treatments. *P < 0.05 and (ns) P > 0.05 by ANOVA, followed by Sidak's
multiple comparisons test. (F) Schematic of the experimental design
and dosing regimen used for testing intratumoral administration
versus contralateral flank administration (draining the axillary
lymph node) of etoposide-treated B16-Ova cells (injured cell
adjuvant) in combination with systemic anti-PD1 and anti-CTLA4. This
was performed independently of the cohorts used in (B) to (E). (G)
Tumor growth curves for mice treated with intratumoral (IT) or
distant lymph node (axillary LN, opposite flank) injection of ex vivo
etoposide-treated B16-Ova cells (injured cell adjuvant) in
combination with systemic anti-PD1 and anti-CTLA4. n indicates the
number of mice in each group. (H) Kaplan-Meier survival curves of the
experiment in (G). *P < 0.05 by log-rank test.
To test whether the site of injection of the injured cell adjuvant
was critical for the subsequent antitumor response in combination
with checkpoint blockade, we compared direct intratumoral injection
with a subcutaneous injection draining into the axillary lymph node
in the opposite flank from that bearing the primary tumor (Fig. 4F).
Mice that received intratumoral injections showed superior tumor
control, survival, and cure compared to animals that received
subcutaneous injections at a distant site (Fig. 4G).

Intratumoral injection of ex vivo etoposide-treated tumor cells
results in persistent cure and immunological antitumor memory in a
subset of treated mice

The subset of mice that demonstrated complete tumor regression after
injured cell adjuvant treatment remained tumor-free for at least 98
days (Fig. 4C). These complete responders and naive control mice
(which were never previously exposed to B16-Ova tumor cells) were
rechallenged in the contralateral flank with live B16-Ova cells (Fig.
5A, top). Tumors grew to 200-mm^2 cross-sectional area within 30 days
in the naive mice (Fig. 5A, bottom), at which point they were
euthanized. None of the intratumoral adjuvant-treated animals who
were cured of their initial tumors after therapy developed tumors
upon rechallenge, suggesting that combining systemic checkpoint
blockade with intratumoral injections of the injured cell adjuvant
induces antitumor immunological memory.
[scisignal]
Fig. 5. Long-term immunological memory and CD8^+ T cell responses
toward non-Ova antigens are induced by the injured cell adjuvant plus
ICB.
(A) Top: Schematic of the rechallenge experiment to monitor tumor
growth and a separate rechallenge experiment in an independent cohort
for ELISpot. To monitor tumor growth, five naive mice and five mice
that demonstrated complete tumor regression after treatment with
injured cell adjuvant + systemic anti-PD1/CTLA4 were rechallenged in
the opposite flank with 100,000 live B16-Ova cells. For ELISpot, an
independent cohort of mice that demonstrated complete tumor
regression after treatment with injured cell adjuvant + systemic
anti-PD1/CTLA4 was rechallenged with 100,000 live B16-Ova cells. On
day 6 after rechallenge, spleens were harvested, and splenocytes were
restimulated with B16-F10, B16-Ova cells, or with purified SIINFEKL
peptide. Spleens from naive mice were also restimulated as above.
Bottom: Resulting tumor growth curves from the rechallenge experiment
(on five cured mice derived from three independent experiments) to
monitor tumor growth. Error bars indicate SEM. (B) Top:
Quantification of ELISpot results (from five cured mice derived from
three independent experiments) expressed as IFN-g-producing cells per
10^5 splenocytes. Error bars indicate SEM. ***P < 0.005 by two-way
ANOVA. Bottom: Representative images from the ELISpot plate for the
conditions in the top.
To examine whether this response was unique to the B16 cell line or
to cells engineered to express the ovalbumin antigen, we performed
similar intratumoral injections of saline- or etoposide-treated tumor
cells, in the presence or absence of systemic ICB, with MC-38 murine
colon carcinoma cells that do not express ovalbumin (fig. S4A). In
this MC-38 tumor model, there was minimal benefit of ICB alone when
the tumors were injected with saline (fig. S4, A and B). Similarly,
direct intratumoral injection of etoposide-treated MC-38 tumor cells
into preexisting MC-38 tumors failed to elicit an antitumor immune
response in the absence of systemic ICB. However, 20% of the animals
who received the combination of the etoposide-treated MC-38 tumor
cells together with systemic ICB showed complete tumor regression and
prolonged survival.
Last, to evaluate whether T cell responses were being elicited toward
nonovalbumin endogenously expressed tumor antigens in the B16-Ova
model, we rechallenged mice that showed complete tumor regression
with live B16-Ova cells and euthanized them 6 days later. The spleens
were harvested and the splenocytes then restimulated in vitro with
B16-Ova cells, the parental B16F10 cell line that does not express
ovalbumin, or with the SIINFEKL peptide, and the number of
IFN-g-producing cells was examined using an enzyme-linked immunospot
(ELISpot) assay. All three restimulation conditions resulted in the
appearance of IFN-g-positive cells, whereas no such cells were seen
in the spleens of naive mice that had not been previously inoculated
with B16-Ova cells (Fig. 5B). The appearance of IFN-g-positive cells
upon restimulation with B16F10 cells lacking ovalbumin expression
indicates that T cells in the cured mice can recognize endogenously
expressed tumor antigens in addition to the artificially expressed
ovalbumin protein.

Batf3^-/- mice do not respond to the injured cell adjuvant and
checkpoint blockade combination

To test whether the efficacy of the injured cell adjuvant in
combination with ICB treatment for an antitumor immune response
depends on DCs that can cross-present tumor antigens, we enumerated
the numbers of CD11b^+C103^- DC2 cells and CD11b^-CD103^+ DC1 cells
by immunophenotyping and flow cytometry (fig. S5A). CD11b^-CD103^+
DC1 cells, which are typically also basic leucine zipper ATF-Like
transcription factor 3^+ (BATF3^+) (39, 40), are known to
cross-present tumor antigens to CD8^+ T cells (41). As above,
flank-implanted B16-Ova tumors were injected directly with saline or
the etoposide-treated injured cell adjuvant in the presence or
absence of systemic checkpoint blockade (Fig. 6A). We also included a
cohort of mice that was treated with intratumoral etoposide in
combination with checkpoint blockade. After two doses of the injured
cell adjuvant or etoposide and three doses of checkpoint blockade,
immunophenotyping of the tumors revealed an enhanced number of CD103^
+ DC1 in tumors that were treated with the injured cell adjuvant and
checkpoint blockade, compared to the other groups (Fig. 6B). In
addition, cross sections of tumors treated with the injured cell
adjuvant and checkpoint blockade showed markedly enhanced BATF3
staining by immunohistochemistry (Fig. 6C), indicating the enhanced
presence of BATF3^+ DCs, which was not present in the other treatment
groups. Intratumoral injection of free etoposide combined with
checkpoint blockade did not enhance numbers of CD103^+ DC1,
consistent with the lack of T cell expansion and lack of efficacy
seen in vivo (Fig. 3, A to D) and in vitro (Fig. 3F) with this
treatment.
[scisignal]
Fig. 6. Intratumoral administration of the injured cell adjuvant plus
systemic ICB increases DC1 density in the tumor and requires BATF3
for efficacy.
(A) Schematic of the experimental design and dosing regimen used to
test the effect of intratumoral etoposide-treated B16-Ova cells in
combination with systemic anti-PD-1/CTLA4 on the frequency of
intratumoral DC. (B) Quantification of intratumoral CD11b^-CD103^+
DC1 and CD11b^+CD103^- DC2 subsets from treated tumors analyzed by
flow cytometry (five mice per group). Error bars represent SEM. *P <
0.05 by ANOVA, followed by Sidak's multiple comparisons test. ns
indicates P > 0.05 by ANOVA. (C) Tumor sections were stained with an
antibody to BATF3. Insets show higher-magnification images of the
boxed central region of each section. Scale bar, 400 mm. (D) Tumor
growth curves of Batf3^-/- mice treated with intratumoral saline or
etoposide-treated B16-Ova cells (injured cell adjuvant) in
combination with systemic anti-PD1 and anti-CTLA4 antibodies. n
indicates the number of mice in each group. (E) Kaplan-Meier survival
curves of the experiment in (D). P = 0.5220 by log-rank test. (F)
Frequency of circulating H2-K^b/SIINFEKL-specific CD8^+ T cells from
WT and Batf3^-/- mice treated as indicated. (G) Quantification of
intratumoral CD86^+ DC1 from tumors that were treated 24 hours before
with one dose of saline or injured cell adjuvant (IT) in presence or
absence of one dose of systemic anti-PD-1 and anti-CTLA4 administered
24 hours before saline or injured cell adjuvant. Error bars represent
SEM, five mice per group. *P < 0.05 and **P < 0.01 by ANOVA followed
by Sidak's multiple comparisons test.
To directly validate the contribution of BATF3^+CD11b^-CD103^+ DC1
cells to antitumor immunity induced by the combination of our injured
cell adjuvant and checkpoint blockade, we repeated the experiment
above (Fig. 4A) using Batf3^-/- mice. Intratumoral injection of ex
vivo etoposide-treated tumor cells with systemic ICB failed to induce
tumor control or prolong the lifespan of tumor-bearing mice in the
absence of BATF3 (Fig. 6, D and E). Last, whereas the combination of
the injured cell adjuvant and systemic ICB showed an enhanced trend
in the frequency of circulating H2-K^b/SIINFEKL-reactive CD8^+ T
cells in wild-type (WT) mice, this trend was not observed in
BATF3-deficient mice (Fig. 6F).
To further explore the presence of costimulatory markers on DC1
cells, we treated mice bearing B16-Ova tumors with a single dose of
saline or the etoposide-treated injured cell adjuvant intratumorally,
in the presence or absence of a single systemic dose of anti-PD-1/
anti-CTLA4 antibodies given 24 hours previously. The tumors were then
harvested 24 hours later, disaggregated, stained for markers of DC1,
the costimulation molecules CD86 and OX40L, and the chemokine
receptor CCR7, and quantified by flow cytometry. The combination of
intratumoral injection of etoposide-treated injured cells with
systemic checkpoint blockade resulted in enhanced density of
activated DC1 cells within the tumor as early as 24 hours after a
single dose of the adjuvant (Fig. 6G and fig. S5B). Together, these
data strongly suggest that intratumoral administration of ex vivo
etoposide-treated tumor cells as an injured cell adjuvant, in
combination with systemic checkpoint blockade, promotes BATF3^+
DC-mediated antitumor T cell responses leading to improved survival
and complete tumor regressions in a subset of mice concurrent with
long-term antitumor immunological memory.

Live injured cells, rather than dead cells, are the determinants of
DC-mediated IFN-g induction in T cells

We observed that both etoposide and mitoxantrone, which effectively
induced DC-mediated IFN-g in CD8^+ T cells, induced substantial
amounts of apoptotic and nonapoptotic tumor cell death compared to
drugs that failed to elicit an immune response (fig. S6A).
Doxorubicin also caused similar amounts of cell death but was
immunologically silent. Curiously, at the drug doses used here (in
fig. S6A), the specific doses of mitoxantrone and etoposide that were
maximally effective were not the doses that caused the greatest
amount of total cell death.
To investigate whether the magnitude of T cell IFN-g responses
directly correlated with the number of dead cells present in the
treated tumor cell fractions that were coincubated with BMDCs, we
treated tumor cells with increasing doses of etoposide or
mitoxantrone from 0 to 100 mM. Using the assay described above (in
Fig. 1A), B16-Ova cells treated with increasing doses of etoposide
induced a corresponding increase in the magnitude of IFN-g responses
in T cells (Fig. 7A). However, the proportion of dead cells [annexin
V (AnnV) or 4',6-diamidino-2-phenylindole (DAPI) single or double
positive] present in the treated tumor cell mixture increased to ~30%
at 25 mM etoposide but was largely unchanged with greater doses.
Cells treated with 5 mM mitoxantrone induced the maximum T cell IFN-g
response among the doses tested, while cells treated with higher
doses induced a lower IFN-g response that became undetectable at 25
mM and higher (Fig. 7, B and C). The dead-cell proportion in the
mitoxantrone-treated B16-Ova cell mixture was equivalent between 5
and 10 mM (~50%) and increased to greater than 90% at 25 mM and
higher doses (Fig. 7D). These results indicated that there was no
agreement between the proportion of dead cells induced by etoposide
or mitoxantrone treatment and the DC-mediated IFN-g responses in T
cells, although all the doses that were immunologically active
resulted in some degree of cell death, which was generally <=~50%.
[scisignal]
Fig. 7. Live injured cells, rather than dead cells, are the primary
determinants of DC-mediated T cell IFN-g responses.
(A and B) Quantification from three independent experiments of IFN-g^
+ CD8^+ T cells induced by coculture of BMDCs with B16-Ova cells
treated with etoposide (A) or mitoxantrone (B) at the indicated
concentrations for 24 hours. The first lane (-) indicates the
percentage of IFN-g^+ CD8^+ T cells produced by coculture of BMDCs
and T cells in the absence of B16-Ova cells. Error bars indicate SEM.
*P < 0.05, **P < 0.01, ***P < 0.005, ****P < 0.0001, and (ns) P >
0.05 by ANOVA followed by Sidak's (A and B, left) or Dunnett's (B,
right) multiple comparisons test. (C and D) Quantification from two
or three independent experiments of the proportion of live (AnnV and
DAPI double negative; white bars) and dead (sum total of AnnV and/or
DAPI single or double positive; gray bars) cells after treatment of
B16-Ova cells for 24 hours with etoposide or mitoxantrone as
indicated. Data points are shown with means +- SEM; where n = 2, error
bars indicate range. (E and F) Quantification from three independent
experiments of IFN-g^+ CD8^+ T cells induced by coculture of BMDCs
with the indicated B16-Ova (E and F) or MC-38-Ova (G and H) live cell
(AnnV^-/DAPI^-) and dead cell fractions obtained after treatment with
etoposide (50 mM) or mitoxantrone (10 mM). "-" indicates controls.
BMDC was cocultured with each of the fractions or combinations
thereof for 24 hours before OT-1 CD8^+ T cells were added. Live +
dead, whole treated cell mixture; Live, live cell fraction; Dead,
dead cell fraction; Sup, cell-free supernatant; Dead + Sup, dead
cells combined with cell-free supernatant. Error bars indicate SEM.
***P < 0.005 and ****P < 0.0001 by ANOVA followed by Dunnett's
multiple comparisons test. (I) Schematic of the experimental design
and dosing regimen used to test the effect of intratumoral
administration of etoposide-treated B16-Ova cells in combination with
systemic anti-PD1/CTLA4 on the frequency of intratumoral H2-K^b/
SIINFEKL-specific CD8^+ T cells. Dosing frequency of saline, live, or
dead fractions of ex vivo etoposide-treated B16-Ova cells (IT) and
systemic anti-PD1 and anti-CTLA4 was as shown in Fig. 6A. Tumors were
harvested 2 days after the second dose of injured cell adjuvant. (J)
Frequency of intratumoral H2-K^b/SIINFEKL-specific CD8^+ T cells from
tumors treated with the conditions indicated (four to five mice per
group). Error bars represent SEM. *P < 0.05 by ANOVA followed by
Sidak's multiple comparisons test.
To further investigate the specific contribution of the dead and live
fractions of tumor cells induced by chemotherapy treatment, we
fractionated the etoposide- and mitoxantrone-treated cell cultures
into cell-free supernatants dead cell-enriched fractions (AnnV^+ and/
or DAPI^+), or live cell-enriched fractions (AnnV and DAPI double
negative) (fig. S6, B to E). Each fraction was then cocultured with
BMDCs for 24 hours, followed by the addition of OT-1 CD8^+ T cells
for an additional 12 to 15 hours, as described above (for Fig. 1A).
Neither the cell-free supernatants nor the dead cell-enriched
fraction, alone or in combination, was capable of inducing
DC-mediated T cell IFN-g responses (Fig. 7, E to H). Furthermore,
consistent with the finding that cell-free supernatants alone did not
induce T cell priming for IFN-g production, analysis of the cytokines
in the supernatants after coculture of chemotherapy-treated cells
with DCs using a 44-cytokine panel revealed that no single cytokine
was sufficiently changed to explain the differences observed between
the immunogenicity of the different chemotherapy drugs used (fig.
S7A). Cotreatment of tumor cells with brefeldin A in combination with
etoposide or mitoxantrone resulted in only a small reduction in T
cell IFN-g production by the etoposide-treated tumor cells and no
reduction with mitoxantrone treatment (fig. S7B), suggesting that
active secretion of factors from drug-treated tumor cells plays, at
most, a limited role. Lysates generated by subjecting the
chemotherapy-treated total cell mixture to three rounds of
freeze-thawing (between liquid nitrogen and 37degC), upon coincubation
with BMDCs, also failed to induce IFN-g in T cells. In marked
contrast, fractions containing the adherent live injured cells were
the most effective at inducing the expression of IFN-g in OT-1
T-cells. Similar behavior was also noted in the MC-38-Ova cells (Fig.
7, G and H).
Last, to directly verify these effects in vivo, we repeated the
above-described experiments (in Fig. 4A) in which mice bearing
B16-Ova tumors were given intratumoral injections of either the live
injured cell fraction or the dead cell fraction of B16-Ova cells that
had been treated ex vivo with 50 mM etoposide, followed by systemic
administration of a-PD-1/CTLA4 immune checkpoint inhibitors (Fig. 7I
). Two days after the second intratumoral dose of the live injured or
the dead cell fractions, mice were euthanized, and the percentage of
H2-K^b-SINFEKL-specific T cells within the tumor was enumerated using
flow cytometry. Only intratumoral injection of the live injured cell
fraction after chemotherapy treatment resulted in significant
tumor-specific T cell infiltration in vivo (Fig. 7J).
We were perplexed by the failure of the 10 or 50 mM doxorubicin
treatments to induce an effective immunogenic response in our assay,
because this drug has previously reported to be capable of inducing
antitumor immunity in vivo (24, 42). We therefore repeated these
DC-mediated T cell stimulation assays after treatment of the B16-Ova
tumor cells with lower doses of doxorubicin. When the doxorubicin
dose was reduced, substantial IFN-g production could be observed,
with a maximal effect seen at 1 mM (Fig. 8A). Consistent with what we
observed with 50 mM etoposide and 10 mM mitoxantrone, treatment of
the B16-Ova cells with 1 mM doxorubicin also resulted in substantial
DC-mediated T cell proliferation (fig. S1E), in contrast to what we
had observed with the 10 mM dose.
[scisignal]
Fig. 8. Live injured chemotherapy-treated tumor cells require
DNA-damage signaling for the DC-mediated T cell IFN-g response.
(A) B16-Ova cells were treated with doxorubicin at 0.5, 1, 2.5, or 5
mM for 24 hours, washed, and incubated with primary bone
marrow-derived dendritic cells (BMDCs) for another 24 hours. After
this, OT-1 CD8^+ T cells were added and evaluated for intracellular
IFN-g 15 hours later. Quantification of IFN-g^+CD8^+ T cells from
three independent experiments is shown. Error bars represent SEM.
****P < 0.0001 by ANOVA, followed by Dunnett's multiple comparisons
test. (B) B16-Ova cells were treated with doxorubicin, etoposide, or
mitoxantrone at the doses indicated for 24 hours and the live
(attached) fractions were analyzed by fluorescence microscopy after
staining with DAPI (blue channel) and anti-gH2AX (red channel).
Representative images are shown. Scale bar, 20 mm. (C) Quantification
of gH2AX foci intensity from the experiment in (B) is shown. n >= 200
cells per condition. Width in violin plot indicates frequency for
each observed value from maximum to minimum, with dotted line
indicating median. ****P < 0.0001 by ANOVA followed by Dunnett's
multiple comparisons test compared to the DMSO control. (D) B16-Ova
cells were treated with doxorubicin, etoposide, or mitoxantrone at
the doses indicated for 24 hours, followed by separation of the live
and dead fractions and analysis of respective lysates by Western
blotting. Blots are representative of two independent experiments. (E
) Quantification of IFN-g^+CD8^+ T cells (from three independent
experiments) induced by BMDCs after incubation with B16-Ova cells
that were cotreated with 50 mM etoposide and 10 mM ATM kinase
inhibitor KU-55933, ATR kinase inhibitor AZD6738, or DNA-PK inhibitor
NU7441. First lane (-) defined as in Fig. 1C. Error bars represent
SEM. **P < 0.01 and ****P < 0.0001 by ANOVA, followed by Dunnett's
multiple comparisons test versus the "Etop 50 mM" group.

DNA damage signaling in the live-cell fraction is critical for the T
cell IFN-g response

To further investigate the role of DNA damage signaling in the
immunogenicity induced by chemotherapy treatment in the live cell
fraction, we performed immunofluorescence microscopy and validated
that treatment with the maximally effective doses of doxorubicin,
etopiside, and mitoxantrone resulted in intense gH2AX nuclear foci
formation in the live B16-Ova tumor cells (Fig. 8, B and C). Next, a
total population of drug treated cells was separated into live and
dead cell fractions, and lysates of each fraction were immunoblotted
using phospho-specific antibodies directed at the phospho-SQ motif
present on substrates of the DNA damage kinases ataxia-telangiectasia
mutated (ATM), ataxia-telangiectasia and Rad3-related (ATR), and
DNA-dependent protein kinase (DNA-PK) (43). The extent of DNA damage
signaling was more pronounced in the live injured cell fraction (Fig.
8D). Last, to investigate the extent to which this DNA damage
signaling contributed to the immunogenicity phenotype, the cells were
treated with small-molecule inhibitors of ATM, ATR, or DNA-PK before
treatment with 50 mM etoposide and assayed for DC-mediated T cell
production of IFN-g (as described above in Fig. 1A). Pretreatment
with the DNA-PK inhibitor was particularly effective in suppressing
DC-mediated T cell stimulation after chemotherapy treatment, with
modest effects also seen after ATR inhibition (Fig. 8E). Some
reduction in production was also observed upon ATM inhibition,
although this failed to reach statistical significance. The effect
was independent of direct sensing of cytosolic DNA by BMDCs, because
Sting^-/- BMDCs were equally effective in inducing T cell responses
by DNA-damaged tumor cells (fig. S7C). Together with the results
showing that the immunogenicity resided in the live cells (Fig. 7),
these data indicate that DNA damage signaling within the live injured
cell fraction after treatment with genotoxic drugs is critical for
the development of effective antitumor T cell responses.

DISCUSSION

The property of certain chemotherapeutic agents to activate the
immune response has classically been attributed to their ability to
induce immunogenic cell death in cancer cells, as reflected by
multiple markers including calreticulin, HMGB1, and ATP (24, 42, 44,
45). Separately, necroptotic death driven by RIPK3 in tumor cells has
been shown to be effective in inducing an antitumor CD8^+ T cell
response (46, 47). Last, elegant work from Tait's group (48, 49)
using BH3 mimetics has implicated caspase-independent cell death by
mitochondrial outer membrane permeabilization in engaging potent
antitumor immune responses. Each of these studies clearly indicates
the ability of specific types of cell death to stimulate an immune
response, but the role of live stressed and injured cells, rather
than dead cells, in promoting immune activation is not clear.
In this study, we used a novel adaptation of an antigen
cross-presentation assay (50) to identify specific drug- and
dose-dependent DNA-damaging conditions and observed that live
genotoxically injured tumor cells, when cocultured with BMDCs, could
drive T cell activation. Administration of ex vivo
chemotherapy-treated live injured tumor cells directly into a
preexisting tumor in vivo functioned as a potent immune adjuvant when
given in combination with systemic ICB in mouse cancer models. This
treatment led an expansion of CD103^+ intratumoral DCs and an
increase in the frequency of H2-K^b/SIINFEKL-reactive antitumor CD8^+
T cells in the tumor and in the circulation, resulting in markedly
enhanced tumor control and survival compared to injection of an
injured cell adjuvant into the contralateral flank in combination
with ICB or by ICB alone. A subset of mice showed complete tumor
regressions and resistance to rechallenge with live tumor cells,
demonstrating immunological memory. A similar complete regression
response was observed using MC-38 cells lacking ovalbumin, indicating
that the results were not limited to one tumor cell type or to cells
that express a foreign nontumor antigen. The findings are summarized
in a model figure (Fig. 9).
[scisignal]
Fig. 9. Model of immunogenic cell injury generating antitumor
immunity.
Intratumoral injection of tumor cells treated ex vivo with
DNA-damaging chemotherapy promotes effective DC-mediated T cell
priming and expansion when combined with systemic immune checkpoint
blockade. Neither cell-free supernatants, the dead cell fraction, nor
lysates of the chemotherapy-treated tumor cells generated by three
cycles of freeze-thawing, when coincubated with DC, promoted a T cell
IFN-g response, indicating that some type of active cellular process
beyond cytokine secretion from genotoxically injured tumor cells is
likely involved. Furthermore, if the DCs and/or T cells were also
exposed to the cytotoxic chemotherapy drug, then T cell IFN-g
responses were impaired. Consequently, direct intratumoral injection
of the free chemotherapeutic drug, in combination with systemic ICB
administration, was largely ineffective at inducing an antitumor
immunogenic response.
The most effective drugs at inducing this BMDC-mediated T cell IFN-g
response were topoisomerase II inhibitors when used at doses that
induced some degree of cell death, although the immunogenic activity
resided in the live injured cell fraction. Although calreticulin
appeared to play an important role, neither the release of HMGB1 or
ATP from damaged or dead cells appeared to be involved, suggesting
that the immunogenic determinants in tumor cells after genotoxic
stress reside within or on the surface of the injured cells. The
function of calreticulin in the injured tumor cells awaits further
clarification but could involve mechanisms distinct from its effects
on dead cell uptake by DCs (24), possibly involving its ER-chaperone
activity (51-53). We cannot, however, completely exclude a
contribution from chemotherapy-induced tumor cell death, given that
some of the live injured cells after chemotherapy treatment may die
during the coincubation period with BMDCs.
Both DNA damage signaling and activation of the NF-kB pathway in the
live injured tumor cells was critical for inducing antitumor
immunogenicity. DNA damage signaling is known to activate NF-kB and
contributes to survival of tumor cells after treatment with DNA
damaging drugs (54-57). This process involves NF-kB essential
modulator (NEMO) nuclear import and export, and activation of IkB
kinase by RIPK1 and the p38 MAPK kinase kinase TAK1 (transforming
growth factor b-activated kinase 1) in a cytosolic complex (54),
consistent with our findings of important roles for p38 MAPK and
RIPK1. Our findings that p38 MAPK was more critical for
mitoxantrone-induced immunogenicity and that RIPK1 signaling played a
critical role in B16-Ova cells but not MC-38-Ova cells suggest that
other protein complexes may be involved in engaging NF-kB in a drug-
and cell type-dependent manner. Activation of NF-kB has been
extensively studied previously in the context of tumor cell
resistance to anticancer therapies, but in light of our results
implicating live injured cells in the T cell IFN-g response, this
process may also be a physiological mechanism for cells to become
immunogenic after DNA damage. Exactly how the activation of the DNA
damage responsive phosphatidylinositol 3-kinase-like kinases ATM,
ATR, and DNA-PK, as well as p38 MAPK and NF-kB, contributes to the
immunogenicity of the live injured cells remains to be clarified in
future work but could involve, for example, up-regulation of
antigenic determinants on the tumor cells themselves (58).
Despite the known importance of RIPK1 activity in promoting
nonapoptotic cell death (59, 60), we did not observe significant
amounts of necroptotic cell death. In this regard, our findings
complement those of Yatim et al. (46) and Snyder et al. (47) who
independently showed a RIPK1-dependent but necroptosis-independent
effect on the CD8^+ T cell response. In those experiments, Yatim et
al. (46) found that artificial induction of RIPK3 in NIH-3T3 cells,
followed by intradermal injection, induced priming of CD8^+ T cells
in vivo in a DC-dependent manner that required RIPK1 activity in the
RIPK3-induced cells. In addition, NF-kB activity was also required.
Snyder et al. (47) reported that overexpression of a synthetic RIPK3
dimerization construct in NIH-3T3 and B16 tumor cells in vitro,
followed by intratumoral administration of these cells, conferred
significant tumor control and antitumor immunity. Transfection with a
RIPK3 variant that is unable to activate RIPK1 but that is still able
to cause necroptotic cell death failed to confer CD8^+ T cell
expansion or tumor control, demonstrating the importance of a
non-necroptotic RIPK1 activity in this process, consistent with our
finding of a role for RIPK1 signaling in live genotoxically injured
cells.
Our results are in good agreement with recent studies wherein a
subset of intratumoral DCs, characterized by their surface expression
of CD103 in mice and blood dendritic cell antigen-3 (BDCA-3) in
humans, was identified as having unique capabilities of
cross-presenting tumor-associated antigens to CD8^+ T cells and
recruiting T cells to the tumor microenvironment through C-X-C motif
ligand 9 or 10 (CXCL9 or CXCL10) (41, 61, 62). The levels of these
DCs in the tumor microenvironment were shown to correlate with better
overall survival in patients with melanoma receiving immune
checkpoint inhibitors (63), consistent with the importance of these
cells in enhancing antitumor immune responses.
Our finding that chemotherapy-induced live injured cells rather than
dead cells are the major determinant of DC-mediated T cell antitumor
responses is entirely consistent with many of the concepts embodied
in the "danger model" of stress-induced immune activation described
by Matzinger in the 1990s [reviewed in (64)]. The danger model
postulated that the expression of costimulatory molecules on
antigen-presenting cells (APCs) can be triggered by "danger signals,"
or "alarm signals," released by the body's own stressed and injured
cells. Consistent with this model, our findings point toward the
concept of "immunogenic cell injury" rather than immunogenic cell
death, as being critical for promoting antitumor T cell responses
after DNA-damaging chemotherapy. Last, our results suggest a
potential method to translate these findings into clinical use. Tumor
cells from patient tumor biopsies could be used together with primary
patient-derived or allogeneic DCs and CD8^+ T cells to screen for the
optimal chemotherapeutic compounds and doses that induce an
immunogenic cell injury response. These injured cells could then be
reinjected into the same tumor in combination with systemic
checkpoint blockade. Although clinical trials will be required to
test efficacy, this approach has potential for patients whose cancers
are accessible for intratumoral delivery and in whom conventional
treatment options have failed and initial or acquired resistance to
ICB has been observed.

MATERIALS AND METHODS

Reagents, cell lines, and mouse strains

Mouse granulocyte-macrophage colony-stimulating factor (GM-CSF) and
AnnV-fluorescein isothiocyanate (FITC) were purchased from BioLegend.
Interleukin-4 (IL-4) was purchased from Thermo Fisher Scientific.
Antibodies to CD3e (FITC; 145-2C11), to CD8 (APC; 53-6.7), to IFN-g
[phycoerythrin (PE); XMG1.2], to CD45 (BUV395; 30-F11), to CD24 (APC;
M1/69), to Ly6C (BV605; AL-21), to F4/80 (BV711; BM8), to major
histocompatibility complex II (MHCII) (PE-Cy7; M5/14.15.2), to CD11b
(BV786; M1/70), to CD103 (BV421; 2E7), to CD86 (clone GL1), to CCR7
(clone 4B12), and to OX40L (clone RM134L) were purchased from
eBioscience or BioLegend. H2-K^b/SIINFEKL tetramer (PE conjugated)
was purchased from MBL Life Science. Nec-1 and Z-VAD were purchased
from Invivogen. Bay 11-7085 and SB202190 were purchased from
Sigma-Aldrich. KU-55933, AZD6738, and NU7441 were from SelleckChem.
Doxorubicin, etoposide, mitoxantrone, cisplatin, paclitaxel,
camptothecin, irinotecan, 5-FU, and cylcophosphamide were purchased
from LC Laboratories or Sigma-Aldrich. Oxaliplatin was purchased from
Tocris Biosciences. Brefeldin A was from BioLegend. An antibody
against ovalbumin was purchased from Abcam (catalog no. ab17293).
Antibodies to Ser^166-phosphorylated RIPK1 (catalog no. 31122S) and
RIPK1 (catalog no. 3493T) were purchased from Cell Signaling
Technology. Antibodies to calreticulin were purchased from Invitrogen
(catalog no. PA3-900) and Cell Signaling Technology (catalog no.
12238T). The phospho-(Ser/Thr) ATM/ATR substrate antibody was
purchased from Cell Signaling Technology (catalog no. 2851S).
CellTiter-Glo was purchased from Promega. CountBright absolute
counting beads for flow cytometry, ammonium-chloride-potassium (ACK)
lysis buffer, Lipofectamine RNAiMAX transfection reagent, LIVE/DEAD
Fixable Aqua Dead Cell Stain kit, and the Cell Trace CFSE
proliferation kit were purchased from Thermo Fisher Scientific. HMGB1
enzyme-linked immunosorbent assay (ELISA) kit was purchased from IBL
International. CD8^+ T cell isolation kit was from STEMCELL
Technologies. Anti-PD1 (clone RMP1-14) and anti-CTLA4 (clone 9D9)
were from Bio X Cell. Antibody to BATF3 was purchased from Abcam (#
ab211304). Antibodies to gH2AX and cleaved poly(adenosine
5'-diphosphate-ribose) polymerase were from Cell Signaling
Technology. B16F10 cells and MC-38 cells were obtained from American
Type Culture Collection. B16F10 cells were engineered to stably
express ovalbumin (B16-Ova cells), as described previously (65).
MC-38-Ova cells were generated by transduction of MC-38 cells with
pLVX-Ova-IRES-hygro, selection of stable expression clones using
hygromycin, followed by isolation and expansion of single-cell
clones. Ovalbumin expression was verified by Western blotting (fig.
S1A). pSIRV-NF-kB-eGFP (enhanced green fluorescent protein) was a
gift from P. Steinberger (Addgene plasmid no. 118093; http://n2t.net/
addgene:118093; RRID: Addgene_118093) (66). To create a B16-Ova NF-kB
reporter line, 293T cells were initially transfected with pSIRV-
[NF-kB-RE]-eGFP and Gag-Pol to generate retrovirus containing
supernatant, which was then used to transduce B16-Ova cells.
Calreticulin siRNA (silencer select ID no. s63272) was purchased from
Thermo Fisher Scientific. C57BL/6J WT, Batf3^-/-, Sting^-/-, and OT-1
mice were purchased from the Jackson Laboratories.

BMDC generation

Bone marrow was harvested from the femurs and tibias of C57BL/6 mice
(Taconic Biosciences). The bone marrow was flushed out after nipping
one end and then centrifuged at 15,000g for 15 s. After one round of
red blood corpuscle (RBC) lysis with ACK lysis buffer, cells were
filtered through a 100-mm filter to remove aggregates, resuspended at
1 x 10^6 cells/ml, and cultured on a 10-cm bacterial plate (12
million cells per plate) in Iscove's modified Dulbecco's medium
(IMDM) containing 10% fetal bovine serum (FBS) with antibiotics, 20
ng/ml each of GM-CSF and IL-4 and 55 mM b-mercaptoethanol. After 3
days, 75% of the medium was replaced with fresh medium containing
growth factors. DCs, which were loosely adherent, were harvested by
gentle pipetting on day 6 or 7, and were used for the assay.

In vitro cross-presentation assay

B16-Ova or MC-38-Ova cells were treated with various doses of
chemotherapeutic drugs for 24 hours, followed by extensive washing in
IMDM [10%FBS with penicillin/streptomycin (P/S)]. Subsequently 1 x 10
^6 treated cells were cocultured with 2.5 x 10^5 BMDCs per well of a
24-well plate for each condition tested. After 24 hours of coculture,
supernatants were removed from each well, and the BMDCs were washed
two to three times in T cell medium (RPMI 1640 containing 10% FBS, 20
mM Hepes, 1 mM sodium pyruvate, 55 mM b-mercaptoethanol, 2 mM l
-glutamine, nonessential amino acids, and antibiotics). CD8^+ OT-1 T
cells isolated from spleens of OT-1 mice were then cocultured with
the BMDCs at 125,000 T cells per well to achieve a T cell-to-BMDC
ratio of 0.5. Where indicated, BMDCs and/or T cells were also exposed
to chemotherapy drugs. After a 12- to 15-hour incubation,
IFN-g-producing T cells were identified and quantified by
intracellular cytokine staining and flow cytometry using a BD LSR II
or LSRFortessa flow cytometer. Cells were first gated for CD3
expression and then regated for CD8 and IFN-g expression. In some
experiments, B16-Ova cells were cotreated with 20 mM Nec-1 or Z-VAD
or 10 mM each of SB202190, Bay 11-7085, KU-55933, AZD6738, or NU-7441
and either etoposide or mitoxantrone at 10 or 50 mM for 24 hours
before performance of the above assay.
For the cross-presentation assay using TC-1 cells to detect expansion
of HPV-E7-specific CD8^+ T cells, TC-1 cells were treated with
various doses of chemotherapeutic drugs for 24 hours, followed by
extensive washing in IMDM (10%FBS with P/S). Subsequently 1 x 10^6
treated cells were cocultured with 2.5 x 10^5 BMDCs per well of a
24-well plate for each condition tested. After 24 hours of coculture,
supernatants were removed from each well, and BMDCs were washed two
to three times in T cell medium as described above. To generate an
antigen-specific T cell population, C57BL/6 mice were injected
subcutaneously at the tail base with 5 nmol of HPV-E7 peptide vaccine
[IDGPAGQAEPDRAHYNIVTF RQIKIWFQNRR(Nle)KWKK] containing 25 mg of
cyclic di-guanosine monophosphate (c-di-GMP) (Invivogen) as an
adjuvant. Primed mice received two booster injections, 14 and 28 days
after the first injection, with the same vaccine formulation. CD8^+ T
cells from spleens of C57BL/6 mice primed and boosted with HPV-E7
peptide vaccine were then isolated on day 6 after the booster dose
and then cocultured with the BMDCs at 125,000 T cells per well to
achieve an effector-to-target ratio of 0.5. After a 12- to 15-hour
incubation, IFN-g-producing T cells were identified and quantified by
intracellular cytokine staining and flow cytometry using a BD LSR II
or LSRFortessa flow cytometer.

CFSE dilution assay

B16-Ova cells were treated with various doses of chemotherapeutic
drugs for 24 hours, followed by extensive washing in IMDM (10% FBS
with P/S). Subsequently, 1 x 10^6 treated cells were cocultured with
2.5 x 10^5 BMDCs per well of a 24-well plate for each condition
tested. After 24 hours of coculture, supernatants were removed from
each well, and the BMDCs were washed two to three times in T cell
medium. CD8^+ OT-1 T cells were isolated from spleens of OT-1 mice
and labeled with CFSE according to the manufacturer's protocol. A
total of 125,000 labeled T cells per well were cultured with BMDCs in
the presence of IL-2 (10 ng/ml) to achieve an effector-to-target
ratio of 0.5. CFSE dilution was assessed by flow cytometric analysis
of the sample on day 5.

Cytokine analysis

B16-Ova cells were treated with various doses of chemotherapeutic
drugs for 24 hours, followed by extensive washing in IMDM (10%FBS
with P/S). Subsequently 1 x 10^6 treated cells were cocultured with
2.5 x 10^5 BMDCs per well of a 24-well plate for each condition
tested. After 24 hours of coculture, supernatants were removed from
each well and sent to Eve Technologies Inc. for the 44-Plex cytokine/
chemokine array (MD44).

Fractionation of live and dead fractions from chemotherapy-treated
cells

B16-Ova cells or MC-38-Ova cells were treated with various doses of
chemotherapy (as indicated in fig. S6) for 24 hours after which the
floating fraction of cells was transferred to a separate tube and
washed with phosphate buffered saline (PBS; for AnnV/DAPI staining)
or IMDM (for coculture with BMDCs). The attached fraction was rinsed
1x with PBS, detached using 5 mM EDTA (in PBS), washed with PBS or
IMDM, and transferred to a separate tube. Separately, cells treated
with chemotherapy for 24 hours were replated at 1 million cells per
well of a 24-well plate in 500 ml of IMDM (10% FBS with P/S).
Cell-free supernatants were collected after a further 24 hours.
Staining with AnnV and DAPI of the attached and floating fractions
after chemotherapy treatment and fractionation revealed that the
attached fraction is predominantly AnnV and DAPI double negative,
suggesting that most cells in this fraction are live injured cells;
on the other hand, the floating fraction (labeled as "suspension" in
fig. S7) consists of cells that predominantly stain positive for AnnV
and/or DAPI, suggesting that the majority of cells in this fraction
are dead cells. Lysate of the total chemotherapy-treated cell mixture
was generated by three rounds of freeze-thawing by alternate
incubations in liquid nitrogen and a 37degC water bath.

Measurement of immunogenic cell death markers

For measurement of calreticulin surface exposure, B16-Ova cells were
treated for 24 hours with various chemotherapy drugs. All attached
and floating cells were harvested and washed in staining buffer [PBS
containing 0.5% bovine serum albumin (BSA)] and incubated with
anti-calreticulin antibodies for 1 hour on ice. Cells were washed
once in staining buffer and then incubated with secondary Alexa Fluor
488-conjugated secondary antibody for 1 hour at room temperature,
washed again, resuspended in staining buffer, and analyzed by flow
cytometry.
For HMGB1 measurement in cell culture medium, B16-Ova cells were
treated for 24 hours with various chemotherapy drugs, medium was
collected, and floating cells were removed by centrifugation at 250g
for 5 min. Cell-free cell culture medium was collected at this time
for the 24-hour time point. For the 48-hour time point, the total
cell fraction was collected after 24 hours of drug treatment, washed
extensively, and replated for a further 24 hours after which the
cell-free cell culture medium was collected. These samples were then
analyzed by ELISA for HMGB1 according to the manufacturer's protocol.
For measurement of ATP levels, cell-free culture medium obtained as
above was analyzed by CellTiter-Glo according to the manufacturer's
protocol. Values were converted to ATP concentrations using a
standard curve generated using pure ATP.

Calreticulin siRNA assay

B16-Ova cells were transfected with calreticulin or control siRNA (30
nM final concentration) using Lipofectamine RNAiMAX according to the
manufacturer's protocol. Forty-eight hours after transfection, cells
were used for the in vitro cross-presentation assay.

Cell death and viability assays

For assessment of cell death, floating and attached cells were
harvested after 24 or 48 hours of treatment with the indicated
chemotherapeutic drugs. Attached cells were detached using 5 mM EDTA
in PBS. The recovered cells were centrifuged at 250g for 5 min,
washed once in PBS containing 0.9 mM Ca^2+ and 0.5 mM Mg^2+, and then
stained with AnnV-FITC for 15 min in annexin-binding buffer at room
temperature according to the manufacturer's protocol (BioLegend).
Cells were costained with DAPI at a final concentration of 1 mg/ml
for 2 min in annexin-binding buffer, brought to a final volume of 500
ml using PBS containing 0.9 mM Ca^2+ and 0.5 mM Mg^2+, and analyzed
by flow cytometry.
For assays of survival, 15,000 cells were plated per well in a
96-well plate in 100 ml of medium with five replicates per condition.
Wells along the four edges of the plate were not used. After cell
attachment, the indicated drugs were added in an equal volume of
medium and incubated for an additional 48 hours. The medium was then
removed and replaced with 100 ml of fresh medium at room temperature.
After a 30-min incubation at room temperature, 50 ml of CellTiter-Glo
reagent was added, followed by 2 min of gentle mixing. The plate was
incubated at room temperature for an additional 10 min. A total of
100 ml of supernatant was transferred to a 96-well white opaque
plate, and luminescence was read on a Tecan microplate reader. Values
were normalized to those of DMSO-treated control cells.

ELISpot assay

Naive mice (that were never injected with tumor cells) and mice that
showed complete tumor regression after treatment with intratumoral
injured cell adjuvant and systemic anti-PD1 and anti-CTLA4 antibodies
were rechallenged with 100,000 B16-Ova cells ipsilateral to the prior
injection. On day 6 after rechallenge, mice were euthanized, and
spleens were harvested. After RBC lysis with ACK lysis buffer,
splenocytes were seeded as the effector cell population at 0.5 x 10^6
cells per well into mouse IFN-g ELISpot plates (BD Biosciences).
Target B16F10 or B16-Ova cells were treated overnight with mouse
IFN-g (500 U/ml) and then g-irradiated at 120 grays. Irradiated tumor
cells were cocultured with splenocytes at 25,000 cells per well for a
period of 24 hours in a 37degC per CO[2] incubator. Plates were
developed according to the manufacturer's protocol and scanned using
the Cellular Technology Limited ImmunoSpot Plate Reader. Quality
control of spot identification was performed using Cellular
Technology Limited ImmunoSpot software by a blinded researcher.

Immunofluorescence

Immunofluorescence was performed by seeding cells onto glass
coverslips in 12-well plates and treated with chemotherapy drugs.
After 24 hours, floating cells were removed, and the attached cells
were washed in PBS and then fixed with a 4% paraformaldehyde solution
for 20 min at room temperature. Cells were blocked in 16.6% goat
serum, 0.3% Triton X-100, 20 mM sodium phosphate, and 0.45 M sodium
chloride before incubation with primary antibody (anti-gH2AX)
overnight, followed by secondary antibodies conjugated with Alexa
Fluor 488 (Invitrogen). Coverslips were then mounted using ProLong
Gold Antifade Mountant (Invitrogen, P36934) and imaged on EVOS FL
Auto Cell Imaging System (Invitrogen). CellProfiler 3.0.0 (developed
by the Broad Institute of MIT and Harvard's Imaging Platform and
available at www.cellprofiler.org) was used to quantify the
integrated intensity of cells stained with gH2AX antibody. Each image
was broken down into its component grayscale images for analysis
using a CellProfiler pipeline. An illumination function module was
used to uniformly reduce and smoothen background staining on the
entire image set. Cell nuclei were defined as primary objects using
the DAPI grayscale images. The integrated intensity within the
primary objects was then quantified for gH2AX foci.

Mouse studies

B16-Ova cells or MC-38 cells (1 x 10^6) were implanted subcutaneously
in the right flank of 7- to 8-week-old female C57BL/6J WT or
BATF3-deficient (Batf3^-/-) mice. After 11 to 13 days, tumors of an
approximate 16-mm^2 median cross-sectional area were typically
detectable by palpation. Mice with tumors were then binned into
groups and injected intratumorally once a week for 3 weeks with 30 ml
of PBS, free etoposide to achieve a final concentration of 50 mM in
the tumor volume, or 1 x 10^6 etoposide-treated cells (24 hours of
drug treatment, followed by extensive washing with PBS). Where
indicated, groups also received intraperitoneal injections of 200 mg
each of anti-PD1 (clone RMP1-14; Bio X Cell) and anti-CTLA4 (clone
9D9; Bio X Cell) twice a week for 3 weeks. Where indicated,
etoposide-treated cells were injected subcutaneously at the base of
the foreleg into the opposite flank draining the axillary lymph node.
Cross-sectional area of tumors was measured in square millimeters
using calipers every 2 to 3 days. In tumor rechallenge experiments,
naive mice controls or mice who had complete tumor regression and
remained tumor free for at least 60 days were subcutaneously injected
in the left flank (contralateral to the initial tumor) with 0.1 x 10^
6 B16-Ova cells, and tumor development was monitored for another 60
days.
To enumerate circulating tumor antigen-specific CD8^+ T cells, mice
were bled retro-orbitally after the second intratumoral dose of PBS,
etoposide, or etoposide-treated tumor cells, and H2-K^b/
SIINFEKL-tetramer-positive CD8^+ T cells analyzed by flow cytometry.
Briefly, 50 ml of whole blood was collected by retro-orbital
bleeding, centrifuged at 250g for 5 min, followed by three rounds of
RBC lysis in 200 ml of ACK buffer. Cells were then washed once in
tetramer stain buffer (PBS containing 5 mM EDTA, 1% BSA, and 50 nM
dasatinib) and stained with PE-conjugated tetramer for 40 min at room
temperature, followed by costaining with antibody to CD8 for 10 min
at 4degC. Cells were then stained with DAPI, washed, and resuspended in
tetramer stain buffer for flow cytometry analysis.

Immunophenotyping

Phenotypic characterization of immune cell populations was performed
by flow cytometry. Briefly, tumors were harvested and mashed through
a 70-mm filter. Collected cells were washed in fluorescence-activated
cell sorting buffer (PBS containing 5 mM EDTA and 1% BSA),
resuspended, and counted. Five million cells from each sample were
stained with fixable Live/Dead Aqua, stained with
fluorophore-conjugated antibodies on ice for 30 min, washed,
resuspended in 450 ml, supplemented with 50 ml of CountBright
absolute counting beads, and analyzed on a BD LSRFortessa flow
cytometer. DCs were scored as CD45^+Ly6C^-CD24^+MHCII^+F480^- (CD11b^
+ or CD103^+) cells using the gating strategy described in (67).

Statistical analysis

All statistical analyses of data were performed using GraphPad Prism
software. Comparisons of multiple experimental treatments to a single
control condition were analyzed by analysis of variance (ANOVA),
followed by Dunnett's multiple comparisons test. Comparisons between
specific treatment groups were analyzed using a Student's t test with
Bonferroni correction for multiple hypothesis testing. For greater
than three specific comparisons, ANOVA followed by Sidak's multiple
comparisons test was used.

Acknowledgments

We thank L. Suarez-Lopez, J. Patterson, B. Van De Kooij, D. Lim, and
C. Barrett for many helpful discussions and the Koch Institute's
Robert A. Swanson (1969) Biotechnology Center for technical support,
including the Flow Cytometry and the Hope Babette Tang (1983)
Histology core facilities.
Funding: This study was supported in part by the NIH (R01-CA226898
and R35-ES028374 to M.B.Y., Cancer Center support grant P30-CA14051,
Center for Environmental Health support grant P30-ES002109), and
awards from the Charles and Marjorie Holloway Foundation, the MIT
Center for Precision Cancer Medicine, and the Ovarian Cancer Research
Foundation to M.B.Y. This study was also supported by the Marble
Center for Cancer Nanomedicine, the Koch Institute Frontier Research
program (to D.J.I.), Mazumdar-Shaw International Oncology Fellowship
to G.S., and the NIH Interdepartmental Biotechnology Training Program
(T32-GM008334 to L.E.M.). D.J.I. is an investigator of the Howard
Hughes Medical Institute.
Author contributions: G.S. and M.B.Y. conceived the study, designed
the experiments, interpreted the results, and wrote the manuscript.
M.B.Y. also obtained funding for the study. D.J.I. interpreted the
results, designed experiments, and supported the study with funding.
G.S. and L.E.M. designed and performed the experiments, analyzed the
data, and interpreted the results. L.E.M. also contributed to writing
the manuscript. J.-K.C. generated a stable cell line for the study
and prepared figures. Y.W.K. and E.D.H. performed specific
experiments. Y.W.K. also analyzed data and prepared figures. W.A.
helped with animal colony maintenance. S.S. helped prepare the model
figure. B.A.J. provided critical input on the statistical tests used.
Competing interests: G.S., L.E.M., D.J.I., and M.B.Y. are inventors
on patent application PCT/US2020/052775 submitted by MIT that covers
the use of live chemotherapy-injured cells as an immune adjuvant for
immunotherapy of cancer. The other authors declare that they have no
competing interests.
Data and materials availability: All data needed to interpret the
study's findings are present in the paper or the Supplementary
Materials.

Supplementary Materials

This PDF file includes:

Figs. S1 to S7
Download (1.12 MB)
View/request a protocol for this paper from Bio-protocol.

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Information & Authors

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Science Signaling
Volume 14 * Issue 705 * 19 October 2021

History

Received: 26 April 2020
Accepted: 23 September 2021
Published online: 19 October 2021

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Copyright (c) 2021 The Authors, some rights reserved; exclusive
licensee American Association for the Advancement of Science. No
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Authors

Affiliations

Ganapathy Sriram^+ https://orcid.org/0000-0002-1651-3607
Department of Biological Engineering, Massachusetts Institute of
Technology, Cambridge, MA 02142, USA.
Department of Biology, Massachusetts Institute of Technology,
Cambridge, MA 02142, USA.
Center for Precision Cancer Medicine, Massachusetts Institute of
Technology, Cambridge, MA 02139, USA.
David. H. Koch Institute for Integrative Cancer Research,
Massachusetts Institute of Technology, Cambridge, MA 02139, USA.
View all articles by this author
Lauren E. Milling^+ https://orcid.org/0000-0002-1053-0887
Department of Biological Engineering, Massachusetts Institute of
Technology, Cambridge, MA 02142, USA.
David. H. Koch Institute for Integrative Cancer Research,
Massachusetts Institute of Technology, Cambridge, MA 02139, USA.
View all articles by this author
Jung-Kuei Chen https://orcid.org/0000-0003-4964-2031
Department of Biological Engineering, Massachusetts Institute of
Technology, Cambridge, MA 02142, USA.
Department of Biology, Massachusetts Institute of Technology,
Cambridge, MA 02142, USA.
Center for Precision Cancer Medicine, Massachusetts Institute of
Technology, Cambridge, MA 02139, USA.
David. H. Koch Institute for Integrative Cancer Research,
Massachusetts Institute of Technology, Cambridge, MA 02139, USA.
View all articles by this author
Yi Wen Kong https://orcid.org/0000-0002-4223-971X
Department of Biological Engineering, Massachusetts Institute of
Technology, Cambridge, MA 02142, USA.
Department of Biology, Massachusetts Institute of Technology,
Cambridge, MA 02142, USA.
Center for Precision Cancer Medicine, Massachusetts Institute of
Technology, Cambridge, MA 02139, USA.
David. H. Koch Institute for Integrative Cancer Research,
Massachusetts Institute of Technology, Cambridge, MA 02139, USA.
View all articles by this author
Brian A. Joughin https://orcid.org/0000-0003-1022-9450
Department of Biological Engineering, Massachusetts Institute of
Technology, Cambridge, MA 02142, USA.
Center for Precision Cancer Medicine, Massachusetts Institute of
Technology, Cambridge, MA 02139, USA.
David. H. Koch Institute for Integrative Cancer Research,
Massachusetts Institute of Technology, Cambridge, MA 02139, USA.
View all articles by this author
Wuhbet Abraham
David. H. Koch Institute for Integrative Cancer Research,
Massachusetts Institute of Technology, Cambridge, MA 02139, USA.
View all articles by this author
Susanne Swartwout
Department of Biological Engineering, Massachusetts Institute of
Technology, Cambridge, MA 02142, USA.
Department of Biology, Massachusetts Institute of Technology,
Cambridge, MA 02142, USA.
Center for Precision Cancer Medicine, Massachusetts Institute of
Technology, Cambridge, MA 02139, USA.
David. H. Koch Institute for Integrative Cancer Research,
Massachusetts Institute of Technology, Cambridge, MA 02139, USA.
View all articles by this author
Erika D. Handly https://orcid.org/0000-0001-8788-3050
Department of Biological Engineering, Massachusetts Institute of
Technology, Cambridge, MA 02142, USA.
Center for Precision Cancer Medicine, Massachusetts Institute of
Technology, Cambridge, MA 02139, USA.
David. H. Koch Institute for Integrative Cancer Research,
Massachusetts Institute of Technology, Cambridge, MA 02139, USA.
View all articles by this author
Darrell J. Irvine^* https://orcid.org/0000-0002-8637-1405
[email protected]
Department of Biological Engineering, Massachusetts Institute of
Technology, Cambridge, MA 02142, USA.
David. H. Koch Institute for Integrative Cancer Research,
Massachusetts Institute of Technology, Cambridge, MA 02139, USA.
Department of Materials Science and Engineering, Massachusetts
Institute of Technology, Cambridge, MA 02139, USA.
Ragon Institute of Massachusetts General Hospital, Massachusetts
Institute of Technology and Harvard University, Cambridge, MA 02139,
USA.
Howard Hughes Medical Institute, Chevy Chase, MD 20815, USA.
View all articles by this author
Michael B. Yaffe^* https://orcid.org/0000-0002-9547-3251
[email protected]
Department of Biological Engineering, Massachusetts Institute of
Technology, Cambridge, MA 02142, USA.
Department of Biology, Massachusetts Institute of Technology,
Cambridge, MA 02142, USA.
Center for Precision Cancer Medicine, Massachusetts Institute of
Technology, Cambridge, MA 02139, USA.
David. H. Koch Institute for Integrative Cancer Research,
Massachusetts Institute of Technology, Cambridge, MA 02139, USA.
Divisions of Acute Care Surgery, Trauma, and Surgical Critical Care
and Surgical Oncology, Department of Surgery, Beth Israel Deaconess
Medical Center, Harvard Medical School, Boston, MA 02215, USA.
View all articles by this author

Notes

*Corresponding author. Email: [email protected] (D.J.I.);
[email protected] (M.B.Y.)
+
These authors contributed equally to this work

Funding Information

http://dx.doi.org/10.13039/100000002National Institutes of Health:
R01-CA226898
http://dx.doi.org/10.13039/100000002National Institutes of Health:
R35-ES028374
http://dx.doi.org/10.13039/100000002National Institutes of Health:
T32-GM008334
http://dx.doi.org/10.13039/100000054National Cancer Institute:
CA226898
http://dx.doi.org/10.13039/100000057National Institute of General
Medical Sciences: T32-GM008334
http://dx.doi.org/10.13039/100000066National Institute of
Environmental Health Sciences: ES028374
http://dx.doi.org/10.13039/100000066National Institute of
Environmental Health Sciences: P30-ES002109
http://dx.doi.org/10.13039/100017397UVA Cancer Center: P30-CA14051
Environmental Health Support: P30-ES002109
http://dx.doi.org/10.13039/100018776Cancer Center, University of
Florida Health: P30-CA14051
Mazumdar-Shaw International Oncology:
Charles and Marjorie Holloway Fund:
MIT Center for Precision Cancer Medicine:
http://dx.doi.org/10.13039/100001282Ovarian Cancer Research Fund:
Marble Center for Cancer Nanomedicine:
Koch Institute Frontier Research Program:

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